Published: June 10th, 2021
We present a protocol to expose the brainstem of adult mouse from the ventral side. By using a gradient-refractive index lens with a miniature microscope, calcium imaging can be used to examine the activity of inferior olive neural somata in vivo.
Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the cerebellum. IO has long been proposed to be crucial for motor control and its activity is currently considered to be at the center of many hypotheses of both motor and cognitive functions of the cerebellum. While its physiology and function have been relatively well studied on single-cell level in vitro, presently there are no reports on the organization of the IO network activity in living animals. This is largely due to the extremely challenging anatomical location of the IO, making it difficult to subject to conventional fluorescent imaging methods, where an optic path must be created through the entire brain located dorsally to the region of interest.
Here we describe an alternative method for obtaining state-of-the-art -level calcium imaging data from the IO network. The method takes advantage of the extreme ventral location of the IO and involves a surgical procedure for inserting a gradient-refractive index (GRIN) lens through the neck viscera to come into contact with the ventral surface of the calcium sensor GCaMP6s-expressing IO in anesthetized mice. A representative calcium imaging recording is shown to demonstrate the feasibility to record IO neuron activity after the surgery. While this is a non-survival surgery and the recordings must be conducted under anesthesia, it avoids damage to life-critical brainstem nuclei and allows conducting large variety of experiments investigating spatiotemporal activity patterns and input integration in the IO. This procedure with modifications could be used for recordings in other, adjacent regions of the ventral brainstem.
The main goal of systems neuroscience is to understand how spatiotemporal activity patterns of neuronal networks contribute to generation of animal behavior. Thus, fluorescent imaging methodology utilizing calcium-sensitive probes has in the past decade become a main tool for examining neuronal network activity in living animals1,2, as it allows visualization of such dynamics across spatial scales ranging from single cells to mesoscale circuitry. In recent years, the common approach where neural circuits in superficial brain structures (such as cerebral or cerebellar cortices) are imaged through a transparent cranial window3 has been complemented with the use of gradient-refractive index (GRIN) lenses4 allowing examination of network dynamics in deep brain structures. Currently-available GRIN lenses allow reaching into structures several millimeters deep, such as the mouse amygdala, hippocampus and basal ganglia5. However, many regions of interest such as various nuclei in the ventral medulla lie significantly deeper, placing them at the extreme of the GRIN lens reach.
Here, we describe how to overcome this difficulty by taking advantage of the relatively easy accessibility of medulla through the ventral aspect of the brain. Using adult mice where the inferior olive (IO), a nucleus in the ventral medulla, has been virally transfected with a calcium sensor GCaMP6s, we describe the surgical steps (modified from the method described originally in Khosrovani et al. 20076) to place a GRIN lens on the ventral surface of the brain of an anesthetized mouse. Using a miniature microscope, we demonstrate the feasibility of recording neuronal activity in such extremely ventral brain regions. While the procedure is necessarily a non-survival surgery and no experimentation can be performed in awake animals, the method allows examination of intact network dynamics in the context of sensory or other afferent pathway stimulation, providing clear advantages over ex vivo-approaches such as using acute slice preparations.
All applicable international, national, and institutional guidelines for the care and use of animals were followed. Aseptic surgery techniques were applied to the stereotaxic virus vector injection.
1. Stereotaxic virus vector injection
NOTE: Virus carrying the genetic material for expressing GCaMP6s (AAV9.CAG.GCaMP6s.WPRE.SV40) is stereotaxically injected as previously described7,8 with following modifications.
Figure 1: Stereotaxic virus vector injection. (a) The laser-pulled quartz glass pipette has a 10-12 mm long straight taper. After pulling, cut off 1-2 mm from the tip. The pipette is finalized by beveling the tip to a 30° needle shape. (b) The correct injection relies on the proper position of mouse body in the stereotaxic frame. Support the mouse chest to prevent stretching the neck. Level the mouse head by aligning the bregma and lambda horizontally. (c) The IO coordination relative to bregma is shown in dorsal view (left) of the mouse skull and coronal view (right top) of the brain. Injection reaches the lateral part of the principal (IOPr) and the dorsal (IOD) subnuclei of IO (right bottom). Please click here to view a larger version of this figure.
2. Preparation of tools and consumables for ventral approach surgery
3. Administration of anesthesia and preparation of the mouse for the surgery
Figure 2: Preparation of ventral approach surgery. (a) Prepare an intubation tube by cutting a 5-6 mm long and 0.8 mm wide slit in the tip of 20-gauge catheter. (b) Mount the animal ventral side up in a stereotaxic frame and adjust the nose cone angle to ensure the animal is breathing easily. Shave the skin around the throat and thigh areas. Attach the SpO2 sensor to the thigh for monitoring mouse vital signs. Insert the rectal temperature probe for monitoring mouse body temperature. Please click here to view a larger version of this figure.
4. Tracheotomy and intubation (20-25 min)
Figure 3: Tracheotomy and intubation of mouse. (a-c) panels show the process of exposing trachea. (a) Remove the throat skin by cutting along the dashed lines. (b) Flip the salivary glands (SG) laterally to expose the trachea covered by the sternothyroid muscle (SM). (c) Slit open SM along the dashed line to expose the trachea. (d-e) panels show the tracheotomy. (d) Support the trachea with a blunt and curved needle. Tie the third trachea ring caudal to the thyroid gland for securing the trachea to the chest skin. (e) Apply isoflurane with an intubation tube with a slit in the tip. Secure the trachea to the chest skin with the suture thread. Secure the intubation tube to the trachea by tying them together. Scale bar in a=5 mm, applies to all panels. Please click here to view a larger version of this figure.
Figure 4: Schematic diagram of ventral approach surgery from lateral view. (a) A schematic drawing with relevant anatomical parts indicated in their relative location when mouse is placed ventral side up. Abbreviations: muscle covering atlas (AM), longitudinal muscle (LM), salivary glands (SG), sternothyroid muscle (SM), thyroid gland (TG). (b) Schematic of arrangement of the intubation tube in relation to the trachea when the tracheotomy is completed. The trachea is secured by the ties on the chest skin (T-trachea). The intubation tube (IT) is secured by the ties around the trachea end (T-tube). (c) Remove the atlas anterior tubercle (AAT) to clear the line of vision to the IO. (d) Schematic describing the positioning of the miniature microscope (MM) and the GRIN lens above the IO for imaging experiment. Please click here to view a larger version of this figure.
5. Exposing the brainstem (40-45 min)
Figure 5: Expose brainstem of mouse for calcium imaging. (a-f) panels show the process of exposing brainstem. (a) Remove the sternothyroid muscle (SM) labeled in Figure 3e. Cut off the larynx and the esophagus. (b) Remove the longitudinal muscle (LM) and the muscle covering atlas (AM). (c) Cut the atlas ventral arches (AVA) with a rongeur and remove the atlas anterior tubercle (AAT). (d) Cut off the occipital bone (OB) to expand the foramen magnum (FM). (e) Expanded FM. (f) The thin cartilage above the foramen magnum is removed. The periosteal layer of dura mater is peeled off. The square indicates the area containing superficial IO neurons. (g) Image the IO with GRIN lens. Scale bar in a=5 mm, applies to a-c. Scale bar in d=2 mm, applies to d-e. Scale bar in f=2 mm. Scale bar in g=2 mm. Please click here to view a larger version of this figure.
6. Calcium imaging
7. Euthanizing animal following procedure
8. Data processing
Here we present a representative recording obtained with the method as described. Figure 6a shows the location of brightly labeled IO cells visualized during the experiment. The dark diagonal stripes are blood vessels. Note the variable brightness of individual cells, resulting from variable transfection efficacy. In panel Figure 6b we show the mean-normalized fluorescence intensity (deltaF/F) traces obtained from the somata indicated with colors and numbers in panel a. Upward deflections represent transient increases in intracellular calcium. Note how different level of GCaMP6s expression (reflected in cell brightness in panel a) lead to variable signal-to-noise-ratios (SNR).
Figure 6: Example recording of activity of IO neurons in anesthetized mouse. (a) Representative example frame from a recording after spatial filtering. Bright spots are IO neuronal somata, several of which have been indicated as regions of interests (ROIs, colored numbers). Dark stripes are blood vessels. (b) Example deltaF/F traces obtained from the ROIs indicated in panel a. Upwards deflections reflect increases in calcium signal. Scale bar in a=10 µm. Please click here to view a larger version of this figure.
As the surgical procedure involves operations performed in the throat region with numerous vitally critical structures (arteries, nerves), it is essential that it is conducted by a researcher with high-level surgical skills. In following, we highlight and comment on several key points of the procedure; however, it must be reminded that no amount of written advice can supplant the experience, skill, and intuition of the researcher.
The most critical step in the surgery is tracheotomy. It involves cutting the trachea, switching isoflurane from the nose cone to the intubation tube, securing the trachea to the chest skin and tying the trachea and the intubation tube together. All these operations must be completed in a smooth and fast manner to avoid accidents, such as inadequate anesthesia, inflow of fluid into trachea or intubation tube slip-off. One must keep the protocol clear in mind before cutting the trachea.
Hemorrhage is one of the major causes of animal death in this surgery. Since the neck area is dense with blood vessels, cut should only be executed when the line of sight is clear to avoid cutting unseen veins and arteries. Therefore, muscles and connective tissues obscuring vision must be removed, and blood from broken capillaries must be cleaned before advancing.
Animal can be kept alive for a long time (more than 8 hours) from the start of surgery. However, it is important to finish the surgical procedure quickly so there is more time to examine brainstem neurons when animal physiological condition is good. A skilled researcher can finish the whole procedure in 70 min.
While the method provides a clean view of ventral brain surfaces, it is unfortunately impossible to do so without performing a tracheotomy as well as removing significant amount of tissue in the throat region. Therefore, the animal cannot be allowed to wake from anesthesia. Furthermore, even though it is possible to keep the animal alive for many hours with careful adjustment of anesthetic delivery, maintaining body temperature and hydration, it is inevitable that prolonged experimentation will eventually lead to weakening of the animal condition. It is left to the expertise of the researcher to consider the maximal duration of stable recordings.
Another potential limitation of the method as described here is that as the GRIN lens is not inserted into the brain parenchyma, only relatively superficial neurons (~150-200 µm) can be examined. While surgical implantation of GRIN lens is technically possible, acute surgery method does not allow sufficient time for neurons to recover from oxidative stress and presence of blood after implantation likely will degrade image quality beyond acceptable.
Despite the above concerns, we believe this is the first time a method for in vivo imaging of IO neurons is presented. It allows examination of spatiotemporal activity in the IO neurons in the context in vivo in the presence of intact afferent inputs from sensory systems as well as the signals from the cerebellar nuclei and the mesodiencephalic junction22, a feat that has not been possible hitherto. With this method, the function of the IO can now be investigated in greater depth with combination of sensory and optogenetic stimulation. Notably, with the evolution of voltage imaging (such as our recent method for voltage imaging in the IO 23), we hope the presented surgical method will inspire numerous researchers to take up the challenge of investigating how the IO contributes to the generation of cerebellar complex spikes.
The authors have nothing to disclose.
We thank Andrew Scott from media center of OIST for his help with video recording and editing. Also, we thank Hugo Hoedemaker for his help with developing the surgery to expose the brainstem and Dr. Kevin Dorgans for his help with drawing diagrams for figures. In addition, great thanks to Salvatore Lacava for his voice-over narration, as well as all nRIM members and pets for continuing support for wellbeing in the tough times of COVID-19.
|Absorbable suture with 6 mm half circle needle
|hook needle with thread
|Castroviejo curved tip needle holder with lock
|Delicate suture tying forceps
|Delicate Suture Tying Forceps
|Dumont #5/45 forceps
|Fine Iris scissors
|Friedman-Pearson rongeur curved tip
|Gelfoam absorbable gelatin sponge
|Hemostatic gelatin sponge
|For virus vector injection
|Graefe Forceps with serrated tip
|It is part of the nVoke2 system. It's designed to nVoke2 miniature microscpe and GRIN lens can be mounted on it
|KETALAR FOR INTRAMUSCULAR INJECTION
|Daiichi Sankyo, Japan
|Laser-Based Micropipette Puller
|Sutter Instrument, USA
|Sutter Instrument, USA
|Motorized Stereotaxic based on Kopf, Model 900
|mouseOxPlus with rectal temperature sensor and thigh clamp pulse oximeter
|Starr Life Sciences, PA, USA
|Measures animal heart rate, arterial oxygen saturation (SpO2), breath rate, and temperature
|nVoke2 integrarted Calcium imaging micro camera system
|Ohaus Compact Scales
|Scale used to weight animal
|Otsuka Normal Saline
|Otsuka Pharmaceutical Factory, Japan
|Physiological-biological temperature controller system
|SuperTech Instruments, Hungary
|Thermal pad for mouse
|ProView Lens Probe 1.0 mm diameter, 9.0 mm length
|Gradient-refractive index (GRIN) lens
|Q114-53-10NP glass capillaries
|Sutter Instrument, USA
|Customized quartz glass capillaries
|Safety IV Catheter 20G
|B. Braun, Germany
|20 gauage catheter used to prepare intubation tube
|Used to polish the tip of 25G needle to prepare curved and blunt needle
|Muromachi Kikai, Japan
|Used to cut the tip of quartz glass pipette
|SomnoSuite low flow inhalation anesthesia system
|Kent Scientific, USA
|Provides precise control of isoflurane flow
|Surgic XT Plus drill
|For virus vector injection
|Syringe 1 ml
|Syringe needle 25G
|Used to make blunt and bended needle
|Syringe needle 26G
|Thrive 2100 Professional Trimmer
|Vannas-Tübingen spring scissors
|Hayashi Pure Chemical, Japan
|Veet sensitive skin
|Hair removal cream
|Xylocaine Jelly 2 % 30ml
|Aspen Japan, Japan
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